Inactivated Whole Virus Particle Influenza Vaccine Induces Anti-Neuraminidase Antibodies That May Contribute to Cross-Protection against Heterologous Virus Infection

Despite the use of vaccines, seasonal influenza remains a risk to public health. We previously proposed the inactivated whole virus particle vaccine (WPV) as an alternative to the widely used split vaccine (SV) for the control of seasonal and pandemic influenza based on the superior priming potency of WPV to that of SV. In this study, we further examined and compared the immunological potency of monovalent WPV and SV of A/California/7/2009 (X-179A) (H1N1) pdm09 (CA/09) to generate immune responses against heterologous viruses, A/Singapore/GP1908/2015 (IVR-180) (H1N1) pdm09 (SG/15), and A/duck/Hokkaido/Vac-3/2007 (H5N1) (DH/07) in mice. Following challenge with a lethal dose of heterologous SG/15, lower virus titer in the lungs and milder weight loss were observed in WPV-vaccinated mice than in SV-vaccinated ones. To investigate the factors responsible for the differences in the protective effect against SG/15, the sera of vaccinated mice were analyzed by hemagglutination-inhibition (HI) and neuraminidase-inhibition (NI) assays to evaluate the antibodies induced against viral hemagglutinin (HA) and neuraminidase (NA), respectively. While the two vaccines induced similar levels of HI antibodies against SG/15 after the second vaccination, only WPV-vaccinated mice induced significantly higher titers of NI antibodies against the strain. Furthermore, given the significant elevation of NI antibody titers against DH/07, an H5N1 avian influenza virus, WPV was also demonstrated to induce NA-inhibiting antibodies that recognize NA of divergent strains. This could be explained by the higher conservation of epitopes of NA among strains than for HA. Taking these findings together, NA-specific antibodies induced by WPV may have contributed to better protection from infection with heterologous influenza virus SG/15, compared with SV. The present results indicate that WPV is an effective vaccine for inducing antibodies against both HA and NA of heterologous viruses and may be a useful vaccine to conquer vaccine strain mismatch.

Introduction
Among pathogens of public health concern are seasonal influenza viruses, which affect over 10% of the world's population and cause considerable morbidity and mortality [1]. Influenza viruses have two surface spike proteins, hemagglutinin (HA) and neuraminidase (NA), which play roles in the attachment to and release from host cells, respectively, and are also major targets of host immunity. Influenza virus RNA-dependent RNA polymerase has high infidelity due to the lack of proofreading activity during virus replication, leading to the accumulation of mutations under the selection pressure of host immunity, which may cause antigenic variation in viral proteins including HA and NA. This phenomenon is referred to as antigenic drift [2][3][4] and leads to the selection of strains that may not be sufficiently neutralized by pre-existing host immunity induced by either infection or vaccination in the past. Therefore, annual vaccinations are recommended to effectively control seasonal influenza, particularly in high-risk groups such as young children and the elderly [5]. Each year, the World Health Organization holds two meetings to select influenza vaccine strains that antigenically match with strains expected to predominate in the forthcoming season in the northern and southern hemispheres based on global epidemiological surveillance data [5][6][7]. Despite these efforts, vaccine efficacy remains low, ranging from 40% to 70%, and is considerably lower when a mismatch occurs, especially in high-risk populations [8][9][10].
Current influenza vaccines are of two types: live attenuated influenza vaccines and inactivated influenza vaccines (IIV) such as split vaccines (SV), whole virus particle vaccines (WPV), and subunit vaccines [11]. SV consists of viral components that are the products of purified virions that are disrupted with ether or detergent, while subunit vaccine undergoes an additional purification step for the further enrichment of HA or NA. Although SV is the most commonly used vaccine due to its low reactogenicity, it poorly primes naïve individuals [12], which is one of the major problems associated with its use. This feature is considered to have resulted from the disruption of virions, leading to poor activation of immunological cascades after vaccination in naïve populations. On the other hand, WPV is the only IIV in which the virus structure is retained, making it the most immunogenic among contemporary IIVs. Previously, we evaluated the immunogenicity of a single inoculation of WPV and SV in mice and macaques. The results indicated that WPV was superior to SV in priming naïve animals and protecting against homologous virus infection [11][12][13], which reproduces the concern with SV in humans. We also showed that the priming potency of WPV may be linked to its ability to significantly activate the maturation of antigen-presenting cells (APCs) through the efficient delivery of viral RNA into the cells [13]. Because of the high potency in priming, WPV should be used as an alternative vaccine, particularly for naïve populations.
When evaluating influenza vaccines, besides the priming potency, the ability to induce cross-immunity needs to be considered. The widely accepted primary assessment of the efficacy of seasonal influenza vaccines is based on how well they induce antibodies against HA, as determined by the hemagglutination-inhibition (HI) assay [2]. An HI titer of ≥1:40 is believed to correlate with a 50% reduction in the risk of contracting severe disease [14][15][16][17]. However, because host immune responses principally target HA, the HA protein is under higher selective pressure than other viral proteins, resulting in a higher rate of antigenic drift [4,18]. Given the importance of antigenic matching between vaccine strains and prevalent strains, the induction of antibodies against other viral proteins such as more antigenically stable NA may be important to overcome the problem of antigenic drift.
In this study, WPV and SV were evaluated for the induction of cross-reactive immunity utilizing a mouse model. For this purpose, we investigated the protection and antibody responses induced by monovalent WPV or SV of A/California/7/2009 (X-179A) (H1N1) pdm09 (CA/09) against an antigenically drifted strain, A/Singapore/GP1908/2015 (IVR-180) (H1N1) pdm09 (SG/15) (https://www.niid.go.jp/niid/en/influenza-e/2382-flu/fluantigen-phylogeny/7810-2018-1-29.html (accessed on 10 May 2022)), and an avian strain A/duck/Hokkaido/Vac-3 (H5N1) (DH/07). Both vaccines similarly induced anti-HA antibody responses against CA/09 and SG/15 after the second vaccination. Importantly, WPV of CA/09 successfully induced anti-NA antibodies against CA/09, SG/15, and DH/07 in mice, while SV did not. Given that the rate of antigenic drift is lower in NA than in HA, the present results indicate the potential of WPV to induce effective cross-immunity against Vaccines 2022, 10, 804 3 of 13 mismatched vaccine strains. The use of WPV might reduce the impact of immune evasion not only by antigenically drifted seasonal influenza virus strains but also by pandemic viruses of subtypes related to contemporary influenza virus strains.

Vaccine Preparation
We prepared monovalent WPV and SV from the same batch of purified virus as previously described [19,20]. Briefly, CA/09 was inoculated in 10-day-old embryonated chicken eggs, which were then incubated at 37 • C for 48 h, and chilled at 4 • C overnight, followed by the harvesting of allantoic fluids. Virus particles were concentrated by highspeed centrifugation at 40,000 g at 4 • C for 90 min, and then purified by centrifugation at 82,000 g at 4 • C for 90 min through a 10-50% sucrose density gradient. The obtained purified virus was divided into two fractions: one fraction was treated with formalin at a final concentration of 0.02% and incubated at 4 • C for 7 days to prepare WPV. To prepare SV, the other fraction was treated with diluted Tween 80 (1:10 ratio with total virus volume; Sigma Aldrich, Burlington, MA, USA) at a final concentration of 0.05% and an equal volume of diethyl ether (Sigma Aldrich, Burlington, MA, USA) and incubated at room temperature for 30 min. After centrifugation at 900 g at room temperature for 15 min, the aqueous phase was collected and diethyl ether was evaporated under nitrogen gas. The protein concentrations of both WPV and SV were measured using a NanoDrop One spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA).

Animals
Eight-week-old female C57BL/6 mice were purchased from Hokudo, Co., Ltd. (Sapporo, Japan), and housed in a BSL-2 animal room at the International Institute for Zoonosis Control under standard laboratory conditions (temperature of 22 • C ± 2 • C, humidity of 50 ± 10%, and a 12 h/12 h light/dark cycle). All mice were given food pellets (CE-2; CLEA Japan, Sapporo, Japan) and water ad libitum.

First Animal Experiment-Vaccination and Virus Challenge Study
Mice were subcutaneously vaccinated twice with either WPV or SV of CA/09 (9 µg protein/100 µL), or PBS as a negative control at a 3-week interval. All vaccinations were performed under inhalation anesthesia with isoflurane (Fujifilm, Wako Pure Chemical Corp., Osaka, Japan). On day 41 from the first vaccination, mice were challenged with a lethal dose of 3000 plaque-forming units (PFU) of SG/15 under the same anesthetic conditions. Daily body weight was measured after the infection with a pre-determined humane endpoint of 25% weight loss relative to weight at the initial measurement. Mice were euthanized by cervical dislocation following anesthesia with isoflurane, at 3-or 5days post-infection (dpi), and lung samples were collected and homogenized in RPMI anti (RPMI 1640 medium supplemented with 20 µg/mL gentamicin, 100 U/mL penicillin, and 100 µg/mL streptomycin). Following centrifugation of the lung homogenate at 400 g for 5 min, the supernatants were collected and stored at −80 • C until use. The experimental schedule is shown in Figure 1a.

Second Animal Experiment-Vaccination and Serum Collection
Mice were subcutaneously vaccinated three times with either WPV or SV of CA/09 (9 µg protein/100 µL), or PBS as a negative control at 3-week intervals. All vaccinations were performed under inhalation anesthesia with isoflurane. Under the same anesthetic conditions, blood was collected from the tail vein on days 19, 41, and 52 from the first vaccination. After clotting, the supernatants were collected as serum samples and stored at −80 • C until use. The serum samples collected on days 19, 41, and 52 were used for the titration of HI, NI, and neutralizing antibodies against CA/09, SG/15, and DH/07 after vaccination with 1, 2, and 3 doses, respectively. The experimental schedule is shown in Figure 2a.

HI Assay
HI assay was performed as previously described [13]. Briefly, collected serum samples were first treated with a three-fold volume of receptor-destroying enzyme (RDE II; Denka Seiken, Tokyo, Japan) to inactivate nonspecific inhibitors and incubated at 37 • C for 15 h, followed by inactivation of RDE by heating at 56 • C for 60 min. PBS was further added to the samples to dilute the initial serum volume by 10-fold. To carry out HI assay, serial two-fold dilutions of RDE-treated sera were performed in 96-well microplates and mixed with 8 hemagglutination units of the virus antigens (CA/09 or SG/15). The plates were incubated at room temperature for 30 min for antibody-antigen reaction. An equal volume of 0.5% chicken red blood cells was added to the mixtures and further incubated for 30 min. The HI antibody titer was determined as the reciprocal of the highest dilution that inhibited hemagglutination.

Neuraminidase and NI Assays
The neuraminidase assay (NA assay) was used to determine the optimal virus concentration for the NI assay. The NA and NI assays described by Aymard-Henry et al. [21] were performed with modifications as follows. For the NA assay, 10 µL of virus stock solution (CA/09/, SG/15, or DH/07) was serially diluted two-fold with PBS in Eppendorf tubes and 20 µL of fetuin substrate (containing 2.5% (w/v) fetuin; Sigma Aldrich, and phosphate buffer (0.4 M Na 2 HPO 4 and 0.4 M NaH 2 PO 4 ); Kanto Chemical Co., Inc., Tokyo, Japan) was added to all tubes. Following incubation at 37 • C with 5% CO 2 for 16 h, 20 µL of periodate reagent (comprising 4.48% (w/v) NaIO 4 and 62% 14 M H 3 PO 4 ); Kanto Chemical Co., Inc., Tokyo, Japan) was added. The tubes were incubated at room temperature for 20 min followed by the addition of arsenite reagent (comprising 10% (w/v) NaAsO 2 , 7.1% (w/v) Na 2 SO 4 , and 0.3% concentrated H 2 SO 4 ; Sigma Aldrich, Burlington, MA, USA) at ten times the volume of periodate reagent, and the tubes were vortexed thoroughly until the brown color disappeared. Then, 500 µL of thiobarbituric acid reagent (comprising 0.6% (w/v) C 4 H 4 N 2 O 2 S and 7.1% (w/v) Na 2 SO 4 ; Kanto Chemical Co., Inc., Tokyo, Japan) was added. Reaction tubes were incubated in boiling water (100 • C) for 15 min. After cooling at room temperature, 600 µL of butanol reagent (containing 95% C 4 H 10 OH and 5% 12 M HCl; Sigma Aldrich, Burlington, MA, USA) was added. The tubes were centrifuged for 10 min at 1600 g, after which 100 µL of the pink chromophore in the butanol phase from each tube was transferred to a transparent flat-bottomed 96-well plate, and the optical density (OD) was measured at 540 nm using an iMark microplate reader (Bio-Rad, Hercules, CA, USA). The virus dilution giving an OD reading of 0.5 was used for the subsequent NI assay.
To perform the NI assay, 10 µL serum samples were serially diluted two-fold with PBS in Eppendorf tubes and equal volumes of diluted virus solutions (as determined by NA assay) were added to each tube. The tubes were incubated at room temperature for Vaccines 2022, 10, 804 5 of 13 60 min, after which 20 µL of fetuin substrate was added, followed by incubation at 37 • C with 5% CO 2 for 16 h. The NA assay protocol was followed for the remaining NI assay steps. The OD was plotted against the sample dilution, and the NI antibody titer was determined as the reciprocal of the highest dilution with ≤50% OD obtained in the control tubes containing PBS instead of serum.

Neutralization of Viral Infectivity Test
Neutralization of viral infectivity test was performed to measure neutralizing antibodies in the serum. A monolayer of MDCK cells was incubated at 37 • C and 5% CO 2 overnight in six-well plates at a density of 4.75 × 10 6 in 3 mL of RP10. The RDE-treated sera prepared as described above were serially diluted two-fold in PBS in a 96-well plate and incubated with 100 PFU of the virus antigens (CA/09 or SG/15) for antigen-antibody reaction. After 60 min, the mixture of virus and serum was added to MDCK cells and incubated at 37 • C and 5% CO 2 for 45 min with shaking every 15 min to allow the non-neutralized virus to adsorb to the cells. Warmed L15 overlay medium (consisting of Leibovitz L-15 (Life Technologies Corp., Carlsbad, NY, USA) with glutamine at pH 6.8 supplemented with 100 U/mL penicillin, 100 mg/mL streptomycin, 0.028% (w/v) NaHCO 3 , 1 mg/mL TPCK-treated trypsin (Worthington Biochemical Corp., Lakewood, NJ, USA), and 0.9% (w/v) agarose) was added to the six-well plates. The plates were incubated at 37 • C with 5% CO 2 for 3 days. Following manual counting of the plaques, the neutralizing antibody titer was determined as the reciprocal of the highest dilution that prevented the growth of plaques to 50% of that obtained in the control wells.

Plaque Assay
A monolayer of MDCK cells was incubated at 37 • C with 5% CO 2 overnight in six-well plates at a density of 4.75 × 10 6 in 3 mL of RP10. The RP10 medium was aspirated, the cell monolayers were washed with RPMI anti , and 125 µL of diluted lung homogenate (10 −1 to 10 −3 ) in RPMI anti was added to each well, except for the control plate. The plates were placed in an incubator at 37 • C with 5% CO 2 for 45 min with shaking every 15 min to allow virus attachment, followed by the addition of warmed L15 overlay medium to each six-well plate. The plates were incubated at 37 • C and 5% CO 2 for 3 days. The plaques were manually counted following plaque formation and the lung virus titers were determined.

Ethics Statement
All animal experiments were performed under approval (approval number 17-0003) from the Animal Care and Use Committee of Hokkaido University following Fundamental Guidelines for the Proper Conduct of Animal Experiment and Related Activities in Academic Research Institutions under the jurisdiction of the Ministry of Education, Culture, Sports, Science and Technology in Japan.

Statistical Analysis
All statistical analyses were performed using GraphPad Prism 8 software (GraphPad Software, San Diego, CA, USA). Significant differences were denoted by p values < 0.05 using one-way analysis of variance (ANOVA) with multiple comparisons (Tukey's multiplecomparison test for parametric one-way ANOVA or Kruskal-Wallis test for non-parametric one-way ANOVA).

Mice Vaccinated with WPV Exhibited Superior Cross-Protection Compared with Those Vaccinated with SV
We compared the protective potency between WPV and SV in mice against heterologous influenza virus SG/15, one of the drifted strains from the vaccine strain CA/09. Three weeks after two subcutaneous vaccinations with either WPV or SV of CA/09 or PBS as a control, mice were challenged with 3000 PFU of SG/15 ( Figure 1a) and monitored daily for changes in body weight (Figure 1b). After infection, mice inoculated with PBS (control group) showed signs of decreased activity and/or a hunched back and over 25% loss of their body weight, reaching the humane endpoint at 5 dpi. Mice vaccinated with SV showed similar clinical symptoms, but mean body weight loss was only approximately 10%. In contrast, mice immunized with WPV demonstrated negligible weight loss and improved activity compared with either PBS-or SV-inoculated group. Virus titers in the lungs at 3 and 5 dpi were determined by the plaque assay (Figure 1c,d, respectively). At 3 dpi, mice in the control group had the highest virus titers in the lungs of over 10 6 PFU, and SV-vaccinated mice had virus titers of around 10 5 PFU. Meanwhile, three out of five WPV-vaccinated mice had undetectable virus titers, and two other mice also had significantly low titers of virus in the lungs compared with mice inoculated with PBS or SV (Figure 1c). At 5 dpi, the virus titers in the lungs of SV-and WPV-vaccinated mice were even lower than at 3 dpi (Figure 1d). Although the difference did not reach significance, virus titers in the lungs at 5 dpi tended to be lower in the WPV group than in the SV group. Based on these results, WPV appears to be more effective than SV to protect against heterologous virus infection.

Mice Vaccinated with WPV Exhibited Superior Cross-Protection Compared with Those Vaccinated with SV
We compared the protective potency between WPV and SV in mice against heterologous influenza virus SG/15, one of the drifted strains from the vaccine strain CA/09. Three weeks after two subcutaneous vaccinations with either WPV or SV of CA/09 or PBS as a control, mice were challenged with 3000 PFU of SG/15 ( Figure 1a) and monitored daily for changes in body weight (Figure 1b). After infection, mice inoculated with PBS (control group) showed signs of decreased activity and/or a hunched back and over 25% loss of their body weight, reaching the humane endpoint at 5 dpi. Mice vaccinated with SV showed similar clinical symptoms, but mean body weight loss was only approximately 10%. In contrast, mice immunized with WPV demonstrated negligible weight loss and improved activity compared with either PBS-or SV-inoculated group. Virus titers in the lungs at 3 and 5 dpi were determined by the plaque assay (Figure 1c,d, respectively). At 3 dpi, mice in the control group had the highest virus titers in the lungs of over 10 6 PFU, and SV-vaccinated mice had virus titers of around 10 5 PFU. Meanwhile, three out of five WPV-vaccinated mice had undetectable virus titers, and two other mice also had significantly low titers of virus in the lungs compared with mice inoculated with PBS or SV (Figure 1c). At 5 dpi, the virus titers in the lungs of SV-and WPV-vaccinated mice were even lower than at 3 dpi (Figure 1d). Although the difference did not reach significance, virus titers in the lungs at 5 dpi tended to be lower in the WPV group than in the SV group. Based on these results, WPV appears to be more effective than SV to protect against heterologous virus infection.  -c) The circles, squares, and triangles indicate data from mice inoculated with PBS, SV, and WPV, respectively. * p < 0.05, ** p < 0.01, *** p < 0.0005, **** p < 0.0001, one-way ANOVA using a multiple-comparison correction; ns indicates not significant (p > 0.05).

After Multiple Vaccinations, Mice Inoculated with SV Induced HI Antibodies Comparable to Those of Mice Vaccinated with WPV
Next, we investigated the induction of humoral immunity by WPV and SV. Here, mice were given an additional third dose to evaluate how both vaccines boost antibodies against HA and NA. Serum samples were serially collected from the same mice after one, two, and three doses of WPV or SV and were evaluated for the induction of HI antibody Vaccines 2022, 10, 804 7 of 13 titers against homologous (CA/09) and heterologous (SG/15) strains (Figure 2a). Three weeks after the first vaccination (primary response), mice vaccinated with WPV showed significantly higher titers of HI antibodies against both CA/09 and SG/15 than mice inoculated with PBS or SV (Figure 2b,e, respectively). While a significant difference in HI antibody titers against CA/09 after the second vaccination was observed between SV and WPV groups (Figure 2c), those against SG/15 were not significantly different between the groups (Figure 2f). Respective HI antibody titers in WPV-and SV-vaccinated animals reached comparable values against both CA/09 and SG/15 after the third and second vaccinations (Figure 2d,g, respectively). These results imply that both WPV and SV have the potential to induce HI antibodies against the homologous and heterologous strains tested here. Furthermore, given similar HI antibody titers against SG/15 in both vaccine groups after the second vaccination, it indicated that there was no significant contribution of HI antibodies to better cross-protection against the heterologous virus challenge in WPV-vaccinated mice shown in Figure 1.

WPV Induced Better NI Antibodies than SV
In addition to anti-HA antibodies, we further evaluated WPV and SV for the induction of antibodies against NA, the second major surface glycoprotein of influenza viruses. Serum samples collected from mice vaccinated with either WPV or SV as described above were subjected to the NI assay. NI antibody titers against homologous CA/09 in the WPVvaccinated group were significantly increased even after the first vaccination, and the values were further elevated by additional vaccinations (Figure 3a-c). On the other hand, NI antibody titers against CA/09 in SV-vaccinated mice ranged from undetectable to very low at all-time points. Even after the third vaccination, two out of five mice produced no detectable NI antibodies (Figure 3c).
We also evaluated the serum titers of the anti-NA antibodies against heterologous strain SG/15 with an NA amino acid sequence identity of 96% (SG/15 NA; Accession No. EPI849450) compared to that of homologous strain CA/09 (CA/09 NA; Accession No. EPI254042,). Although a single inoculation of WPV significantly but only slightly increased the NI antibody titers against SG/15 (Figure 3d), the titers were significantly elevated by the second and third shots of WPV (Figure 3e,f, respectively). The NI antibody titers against SG/15 in SV-vaccinated mice were below the detectable limit at almost all-time points, as in the PBS control group (Figure 3d-f).
We further examined whether either vaccine effectively induces NI antibodies against H5N1 avian influenza virus strain DH/07 with 90% NA amino acid sequence identity compared to that of CA/09 (DH/07 NA; Accession No. EPI3498). Mice vaccinated with SV did not produce detectable NI antibodies after one or multiple vaccinations (Figure 3g-i). Interestingly, NI antibodies against DH/07 were detectable in the WPV-vaccinated mice even after the first inoculation and increased with subsequent doses, reaching a titer of 71 ± 14 after the third shot. The NI antibody titers in the WPV group were significantly higher than those in PBS-or SV-injected groups at all-time points. These findings suggest that WPV has excellent ability to induce effective NI antibodies against a wide range of viral strains, which may be one of the features that distinguishes WPV from SV. Figure 3. NI antibody titers after vaccination with WPV or SV. C57BL/6 mice were subcutaneously vaccinated three times at 21-day intervals with 9 µg of protein of monovalent CA/09 WPV or SV, as shown in Figure 2a. NI antibody titers against homologous CA/09 (a-c) and heterologous SG/15 (df) and DH/07 (g-i) were evaluated in serum samples collected on days 19 (a,d,g), 41 (b,e,h), and 52 (c,f,i). Dots represent individual values, and the mean values with SEM are indicated by lines (n = 5 per group). The circles, squares, and triangles indicate data from mice inoculated with PBS, SV, and WPV, respectively. * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001, one-way ANOVA using a multiple-comparison correction; ns indicates not significant (p > 0.05).
In addition to HI and NI antibody titers, neutralizing antibody titers were also evaluated in serum samples collected 3 weeks after the second inoculation with WPV or SV. Neutralizing antibody titers against homologous (CA/09) or heterologous (SG/15) strains were induced by both WPV and SV, but the titers against both strains were significantly higher in WPV-vaccinated mice than those induced by SV (Figure 4a,b). Taken together with the results of HI and NI antibody titers, the higher titer of neutralizing antibodies observed in WPV-vaccinated mice may be attributed to the superior inducibility of NI antibodies by WPV. Collectively, the findings of the present study suggest that WPV confers better cross-protective immunity against heterologous virus infection than SV. . NI antibody titers after vaccination with WPV or SV. C57BL/6 mice were subcutaneously vaccinated three times at 21-day intervals with 9 µg of protein of monovalent CA/09 WPV or SV, as shown in Figure 2a. NI antibody titers against homologous CA/09 (a-c) and heterologous SG/15 (d-f) and DH/07 (g-i) were evaluated in serum samples collected on days 19 (a,d,g), 41 (b,e,h), and 52 (c,f,i). Dots represent individual values, and the mean values with SEM are indicated by lines (n = 5 per group). The circles, squares, and triangles indicate data from mice inoculated with PBS, SV, and WPV, respectively. * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001, one-way ANOVA using a multiple-comparison correction; ns indicates not significant (p > 0.05).
In addition to HI and NI antibody titers, neutralizing antibody titers were also evaluated in serum samples collected 3 weeks after the second inoculation with WPV or SV. Neutralizing antibody titers against homologous (CA/09) or heterologous (SG/15) strains were induced by both WPV and SV, but the titers against both strains were significantly higher in WPV-vaccinated mice than those induced by SV (Figure 4a,b). Taken together with the results of HI and NI antibody titers, the higher titer of neutralizing antibodies observed in WPV-vaccinated mice may be attributed to the superior inducibility of NI antibodies by WPV. Collectively, the findings of the present study suggest that WPV confers better cross-protective immunity against heterologous virus infection than SV.  Figure 1a. Serum samples collected on day 41 were evaluated for neutralizing antibody titers against CA/09 (a) and SG/15 (b). The circles, squares, and triangles indicate data from mice inoculated with PBS, SV, and WPV, respectively. * p < 0.05, ** p < 0.01, **** p < 0.0001, oneway ANOVA using a multiple-comparison correction.

Discussion
Here, we investigated the protectivity of mice from heterologous virus infection and antibody induction against homologous and heterologous viruses by WPV and SV of CA/09 in mice. In the challenge experiment with the heterologous virus, SG/15, mice vaccinated with WPV had significantly reduced virus titers in the lung, milder body weight loss, and improved activity than those receiving SV (Figure 1b-d), suggesting a superior preventive effect of WPV against severe disease caused by vaccine-mismatched strains. Although these differences in protectivity were observed between WPV and SV, antibodies against HA of SG/15 were comparable after the second vaccination between mice receiving either vaccine (Figure 2f,g). In contrast to anti-HA antibodies, animals inoculated with WPV had higher antibody titers against NA of the heterologous virus SG/15 as well as homologous strain CA/09, which may explain the higher titers of neutralizing antibody against both strains in WPV-vaccinated mice (Figure 3d-f). Furthermore, anti-NA antibodies induced by WPV reacted against the avian influenza H5N1 strain, DH/07 ( Figure  3g-i). While numerous studies have been published on anti-NA antibodies and their benefits to individuals during infection [22][23][24][25][26][27][28], few have compared SV and WPV for their induction of NA antibodies against heterologous and avian influenza viruses. The results presented here highlight one of the disadvantages of the widely used seasonal influenza vaccines, SV, and we believe this knowledge can be a useful guide for the selection of vaccine strain. Vaccination with WPV may have greater potential than SV to protect against infection with heterologous virus strains, in part due to NA antibodies.
The primary assessment of the efficacy of influenza vaccines is based on the induction of anti-HA antibodies, as determined using the HI assay. Based on this assay, both SV and WPV showed potency in inducing anti-HA antibodies, achieving similar levels after two or three inoculations in this study. Interestingly, despite inducing similar levels of HI antibodies against SG/15 after the second vaccination, following virus challenge, the lung virus titers were significantly lower in the WPV-inoculated group at 3 dpi than in the . Neutralizing antibody titers after vaccination with WPV and SV. C57BL/6 mice were subcutaneously vaccinated twice at 3-week intervals with 9 µg of protein of monovalent CA/09 WPV or SV, as shown in Figure 1a. Serum samples collected on day 41 were evaluated for neutralizing antibody titers against CA/09 (a) and SG/15 (b). The circles, squares, and triangles indicate data from mice inoculated with PBS, SV, and WPV, respectively. * p < 0.05, ** p < 0.01, **** p < 0.0001, one-way ANOVA using a multiple-comparison correction.

Discussion
Here, we investigated the protectivity of mice from heterologous virus infection and antibody induction against homologous and heterologous viruses by WPV and SV of CA/09 in mice. In the challenge experiment with the heterologous virus, SG/15, mice vaccinated with WPV had significantly reduced virus titers in the lung, milder body weight loss, and improved activity than those receiving SV (Figure 1b-d), suggesting a superior preventive effect of WPV against severe disease caused by vaccine-mismatched strains. Although these differences in protectivity were observed between WPV and SV, antibodies against HA of SG/15 were comparable after the second vaccination between mice receiving either vaccine (Figure 2f,g). In contrast to anti-HA antibodies, animals inoculated with WPV had higher antibody titers against NA of the heterologous virus SG/15 as well as homologous strain CA/09, which may explain the higher titers of neutralizing antibody against both strains in WPV-vaccinated mice (Figure 3d-f). Furthermore, anti-NA antibodies induced by WPV reacted against the avian influenza H5N1 strain, DH/07 (Figure 3g-i). While numerous studies have been published on anti-NA antibodies and their benefits to individuals during infection [22][23][24][25][26][27][28], few have compared SV and WPV for their induction of NA antibodies against heterologous and avian influenza viruses. The results presented here highlight one of the disadvantages of the widely used seasonal influenza vaccines, SV, and we believe this knowledge can be a useful guide for the selection of vaccine strain. Vaccination with WPV may have greater potential than SV to protect against infection with heterologous virus strains, in part due to NA antibodies.
The primary assessment of the efficacy of influenza vaccines is based on the induction of anti-HA antibodies, as determined using the HI assay. Based on this assay, both SV and WPV showed potency in inducing anti-HA antibodies, achieving similar levels after two or three inoculations in this study. Interestingly, despite inducing similar levels of HI antibodies against SG/15 after the second vaccination, following virus challenge, the lung virus titers were significantly lower in the WPV-inoculated group at 3 dpi than in the SV-vaccinated group. Reports have been published describing that the suppression of viral growth is associated with the induction of anti-NA antibodies [29,30], whose functionality is similar to NA inhibitors, the main antivirals for the therapeutic treatment of influenza. NA inhibitors and anti-NA antibodies prevent virus budding and spread from infected cells, thus suppressing viral growth, particularly during the early stages of infection. Taken together with previous reports, the anti-NA antibodies induced by WPV observed in this study could have contributed to the additional suppression of viral growth, compared with SV. Therefore, understanding of vaccine efficacy could be improved by incorporating analyses involving antibodies against other viral proteins such as NA, in addition to the HI assay.
Several factors may account for the low or negligible induction of anti-NA antibodies by SV. First, ether-and detergent-disrupted SVs are poor immunogens in mice and humans [31,32] because the viral RNA in the SVs, which affects APC activation, is degraded [13]. Second, the HA protein, which binds to sialic acid receptors, is preferentially endocytosed by APCs over NA or other viral proteins, subsequently inducing antibodies mainly against HA. In WPV, the intact virus structure provides an advantage during antigen presentation as the whole virus particle, together with all other virus proteins, can be taken up and processed by APCs, yielding a broader repertoire of antibodies. In the present study, we found that anti-NA antibodies induced by WPV were reactive even to an avian H5N1 virus strain. The ability of WPV to induce both anti-HA and NA antibodies not solely against the homologous virus but also against the heterologous viruses makes its use advantageous for controlling seasonal influenza caused by antigenic variant viruses.
Historically, pre-existing NA antibodies within populations that cross-react with novel strains have been associated with reduced severity of disease, and this was the case during the 2009 and 1968 pandemics caused by H1N1 and H3N2 strains, respectively [33][34][35]. In line with this concept, the use of WPV as a seasonal influenza vaccine strain may partially contribute to the control of pandemic influenza caused by the H5N1 strain via N1 NA antibodies.
Vaccination for the control of seasonal influenza is clearly fundamental for reducing severe morbidity and mortality. A vast number of studies on the development of immunogenic, cross-reactive, and safe vaccines are ongoing. Our series of studies has shown that WPV is not only an excellent vaccine for naïve populations because of efficient priming, but also has the remarkable advantage of protecting against heterologous virus strains. It also induces a broad range of anti-NA antibodies that inhibit NAs of heterologous strains, which may protect against infection with divergent strains. We hereby propose that WPV is ideal not only for the control of seasonal influenza but also for establishing preparedness for future pandemics.
Institutional Review Board Statement: The animal study protocol was approved by the Ethics Committee of Animal Care and Use of Hokkaido University (protocol code 17-0003, 11/4/2019).


Muscle Histopathological Abnormalities in a Patient With a CCT5 Mutation Predicted to Affect the Apical Domain of the Chaperonin Subunit

Recognition of diseases associated with mutations of the chaperone system genes, e.g., chaperonopathies, is on the rise. Hereditary and clinical aspects are established, but the impact of the mutation on the chaperone molecule and the mechanisms underpinning the tissue abnormalities are not. Here, histological features of skeletal muscle from a patient with a severe, early onset, distal motor neuropathy, carrying a mutation on the CCT5 subunit (MUT) were examined in comparison with normal muscle (CTR). The MUT muscle was considerably modified; atrophy of fibers and disruption of the tissue architecture were prominent, with many fibers in apoptosis. CCT5 was diversely present in the sarcolemma, cytoplasm, and nuclei in MUT and in CTR and was also in the extracellular space; it colocalized with CCT1. In MUT, the signal of myosin appeared slightly increased, and actin slightly decreased as compared with CTR. Desmin was considerably delocalized in MUT, appearing with abnormal patterns and in precipitates. Alpha-B-crystallin and Hsp90 occurred at lower signals in MUT than in CTR muscle, appearing also in precipitates with desmin. The abnormal features in MUT may be the consequence of inactivity, malnutrition, denervation, and failure of protein homeostasis. The latter could be at least in part caused by malfunction of the CCT complex with the mutant CCT5 subunit. This is suggested by the results of the in silico analyses of the mutant CCT5 molecule, which revealed various abnormalities when compared with the wild-type counterpart, mostly affecting the apical domain and potentially impairing chaperoning functions. Thus, analysis of mutated CCT5 in vitro and in vivo is anticipated to provide additional insights on subunit involvement in neuromuscular disorders.

INTRODUCTION
Neuromyopathies constitute a large group of diverse diseases typically associated with genetic variants that have a pathogenic impact on nerves and muscles (Morrison 2016; collection of articles in Front Mol Biosci, 2018-Pathologic Conditions of the Human Nervous and Muscular Systems Associated with Mutant Chaperones: Molecular and Mechanistic Aspects). In many of these diseases there is a mutant gene that encodes a component of the chaperone system (CS), e.g., a chaperone gene such as those that code for the chaperonins Hsp60 (Heat shock protein 60) and CCT (Chaperonin Containing TCP1; also called TRiC, TCP1 Ring Complex), and these disorders are classified as neurochaperonopathies (Scalia et al., 2021). An illustrative example of neurochaperonopathy is the distal sensory mutilating neuropathy associated with a point mutation on the CCT5 subunit described earlier (Bouhouche et al., 2006a;Bouhouche et al., 2006b). More recently, we reported a different genetic variant of CCT5 that was associated with a severe distal motor neuropathy (Antona et al., 2020), which is the object of this article.
The hereditary and clinical aspects of chaperonopathies in general, including the neurochaperonopathies, are well established for the most part (Macario and Conway de Macario, 2005;Rubin et al., 2012;Macario et al., 2013). However, the impact of the pathogenic mutation on the properties and functions of the chaperone molecule has been characterized for only very few chaperonopathies, and still less is known on the associated histopathology and the molecular mechanisms responsible for the tissue abnormalities (Macario and Conway de Macario 2020). The scarcity of data on the histopathological manifestations of chaperonopathies impedes advances in the elucidation of the molecular mechanisms underpinning the severe muscular deficiency observed in patients and this, in turn, stands in the way for developing specific treatments. To remedy this lack of necessary knowledge, we studied the available muscle sample from the patient reported earlier, who suffered from a severe motor disorder and carried a point mutation on the CCT5 subunit (Antona et al., 2020). Here, we define histological abnormalities occurring in striated skeletal muscle from this patient. In our ex vivo investigation, we asked the few questions that seemed most relevant under the circumstances, pertaining to the status of 1) the muscle tissue; 2) the CCT5 subunit and other pertinent components of the CS; and 3) other molecules involved in the maintenance of the muscle structure and function. Consequently, we performed experiments to reveal the general architecture of the muscle and to determine if it presented signs of atrophy and apoptosis. We also ran tests to determine whether the presence and distribution of the CCT5 subunit were altered in comparison with normal muscle. Likewise, we tested for the presence and location of other components of the CS known to play key roles in skeletal muscle, i.e., alpha-B-crystallin and Hsp90. These are in normal muscle predominantly localized to the Z discs and we looked to see if the muscle from the patient showed this characteristic. We also ran experiments to determine if CCT5 colocalized with another member of the CCT complex, CCT1, which would indicate that the mutant subunit is competent to integrate the chaperoning complex and does form it. We investigated levels of proteins that are known (e.g., actin) or suspected substrates for CCT because any abnormality of these proteins would be an indicator of deficient chaperoning by the chaperonin. Lastly, we tested for the presence and distribution of desmin, the muscle molecule with key functions in the maintenance of overall structure and connections between the main components of the muscle fiber, essential for the coordinated functioning of all of them. The impact of the mutation on the molecular properties of the mutant CCT5 protein were investigated in silico by modelling and molecular dynamics simulation.

Muscle Tissues and Microscopy
A lateral gastrocnemius skeletal muscle biopsy from the patient, and a corresponding specimen from a healthy individual, were used. The study was approved by the Ethics Committee of University Hospital AUOP Paolo Giaccone of Palermo. Skeletal muscle tissue samples were fixed in 10% buffered formalin and embedded in paraffin. Thin sections (5 µm), obtained from paraffin blocks by microtome, were stained with haematoxylin-eosin and Alcian Blue Pas for histological evaluation. Examination of the sections was performed by two expert pathologists (F. C, and F. R.) with an optical microscope (Microscope Axioscope 5/7 KMAT, Carl Zeiss, Milan, Italy) connected to a digital camera (Microscopy Camera Axiocam 208 color, Carl Zeiss, Milan, Italy).

Immunohistochemistry for Desmin
Sections were obtained from the paraffin blocks of skeletal muscle tissue and dewaxed in xylene for 30 min at 60°C and, after being passed through a descending scale of alcohol concentrations, they were rehydrated in distilled water at 22°C. Subsequently, antigen unmasking was performed with sodium citrate buffer (pH 6) at 75°C for 8 min and then immersion in acetone at −20°C for 8 min. Then, the sections were immunostained using a Histostain ® -Plus third Gen IHC Detection Kit (Life Technologies,073). The primary antibody used was anti-desmin mouse monoclonal diluted 1:50 (Table 1). Appropriate negative controls were run concurrently for each reaction. Nuclear counterstaining was performed using hematoxylin (Hematoxylin aqueous formula, REF 05-06012/LN. Cat. No. S2020, Bio-Optica, Milan, Italy). Finally, the sections were examined with an optical microscope as described in the previous Section by two independent observers (F. C, and F. R.), who evaluated the reactions and the immunolocalization on two separate occasions.

Immunofluorescence
Deparaffinized sections were incubated in the antigenunmasking solution (10 mM tri-sodium citrate, 0.05% Tween-20) for 10 min at 75°C, and treated with blocking solution (3% BSA in PBS) for 30 min. Next, the primary antibody (Table 1), was applied, and the sections were incubated in a humidified chamber overnight at 4°C. Then, the sections were incubated for 1 h at 22°C with a conjugated secondary antibody (anti-rabbit IgG-FITC antibody produced in goat, F0382, Sigma-Aldrich; anti-mouse IgG-TRITC antibody produced in goat, T5393, Sigma-Aldrich). Nuclei were stained with Hoescht Stain Solution (1:1,000, Hoechst 33258, Sigma-Aldrich). The slices on the slides were treated with Perma Fluor Mountant (Thermo Fisher Scientific, Inc. Waltham, MA, United States) and covered with a coverslip. The images were captured using a Leica Confocal Microscope TCS SP8 (Leica Microsystems).

Double Immunofluorescence
Double immunofluorescence was performed as previously described (Marino Gammazza et al., 2014). Briefly, slices of the muscle tissues were deparaffinized, incubated in the antigen unmasking solution for 10 min at 75°C, and treated with blocking solution for 30 min. The sections were then incubated with the first primary antibody ( Table 1) overnight at 4°C. The day after, the sections were incubated with the second primary antibody ( Table 1) overnight in a humidified chamber at 4°C. Afterwards, the tissue slices were incubated with fluorescent secondary mouse IgG antibody TRITC conjugated (Sigma-Aldrich), diluted 1:250, for 1 h at 22°C, and with FITC conjugated rabbit IgG secondary antibody (Sigma-Aldrich), diluted 1:250, for 1 h at 22°C in a moist chamber. The nuclei were counterstained with Hoechst 33342 for 15 min at 22°C. Finally, the slices were covered with two drops of PBS and mounted with coverslips, using a drop of Vectashield. Imaging was immediately performed with a Leica Confocal Microscope TCS SP8 and the fibers' positivity for both markers ("merge") was assessed using the Leica application suite advanced fluorescences software.

In Silico Analyses
We used the structure of crystallized protein deposited in the Protein Data Bank with accession codes 5UYZ (Pereira et al., 2017) to compare the structure of wild type and mutant CCT5 subunit. Study and alignment of CCT5 wild type and mutant linear amino acid sequences was conducted using Clustal Omega multiple sequence alignment program.

Molecular Modelling and Dynamic Simulations
The molecular modelling of the mutant CCT5 subunit was obtained by replacing the amino acid residue Leucine in position 224 with a Valine, using the package Maestro Schrödinger LLC, New York, NY, 2018, version 11.6.010] and compared with the model of the wild type molecule. Molecular Dynamics simulations were performed for 150 ns (in some cases were extended to 200 ns), using the GROMACS 5.1.1 package (Van Der Spoel et al., 2005;Hess et al., 2008). Interactions were described using an all-atoms CHARMM27 force field (Foloppe and MacKerell, 2000;MacKerell and Banavali, 2000). All the simulations for the various systems, and radius of gyration and RMSD analyses were performed as previously described (Antona et al., 2020).

Atrophy of Muscle Fibers and Disruption of the Tissue Architecture
Histological analysis showed changes in muscle from the patient (henceforth designated MUT muscle) compared to muscle from the healthy control (henceforth designated CTR muscle). MUT muscle showed widespread atrophy, hypereosinophilia, and disruption of the tissue architecture (Figures 1A-C; Supplementary Figurs S1). The MUT muscle fibers were different in shape and size from those in CTR muscle being predominantly small and rounded in cross section ( Figures 1A, B). The same features were revealed with Alcian-Pas staining (Supplementary Figurs S1). In longitudinal sections the MUT muscle fibers had a wavy shape ( Figure 1C). In MUT muscle, nuclei were swelled and in contact with the sarcolemma but in some fibers the nuclei were internal, paracentric.

Apoptosis of Muscle Fibers
MUT muscle showed many more apoptotic fibers than CTR muscle by the TUNEL assay ( Figure 2).

CCT5 Presence, Distribution, and Colocalization With CCT1
In the CTR muscle, CCT5 was present in the sarcolemma, cytoplasm, and within the nuclei (Figure 3, CTR panels) but its signal was slighter in the cytoplasm, nuclei, and, but not as much, in the sarcolemma of MUT muscle fibers ( Figure 3, MUT panels). In the MUT muscle, CCT5 was detected in the extracellular space (intercellular matrix) ( Figure 3, MUT panels).
To determine if the CCT5 subunit in the MUT muscle was associated to other components of its chaperoning team, i.e., the CCT oligomer, colocalization experiments were performed, using double immunofluorescence with antibodies against CCT5 and CCT1. The two subunits colocalized in the MUT and CTR muscles ( Figure 4). The results for CCT5 confirm the localization shown in Figure 3: it appears, here together with CCT1, in the sarcolemma, cytoplasm and nuclei of CTR muscle, whereas in the MUT muscle the two subunits appear mostly in FIGURE 1 | Haematoxylin-eosin staining revealed striking differences between the MUT (panels A-C) and CTR (panel D) muscles. In cross-section the MUT muscle fibers appeared rounded, of diverse sizes, with swelled nuclei in contact with the sarcolemma or near it (examples shown by arrows on panels A,B). These fibers in longitudinal section showed a wavy shape with a striated pattern inside (examples shown by arrows on panel C). The inter-fiber space is considerably wider in the MUT than in the CTR muscle. Bar = 100 µm.
FIGURE 2 | Representative image of TUNEL assay of MUT and CTR muscles. DNA-strand breaks (green spots were more abundant in the MUT (white arrows in the panel with enlargement to the far right, bottom) an in the CTR muscle. CN, negative control for MUT and CTR muscles without TdT enzyme. Bar = 100 μm.
Frontiers in Molecular Biosciences | www.frontiersin.org June 2022 | Volume 9 | Article 887336 4 sarcolemma and the extracellular matrix ( Figure 4). The lower signal of CCT1 in nuclei of MUT muscle closely parallel the low signal of CCT5, suggesting these subunits tend to associate with each other in MUT muscle as well.

Actin and Myosin
Actin and myosin distribution patterns in MUT and CTR muscles were similar, but myosin signal was slightly increased, and that of actin was decreased in MUT as compared with CTR muscle ( Figure 5).

Desmin
Desmin (muscle-specific type III intermediate filament) is a muscle-specific protein, normally localized in the sarcolemma, Z-disc, and nuclear membrane, that maintains the sarcomere structure, linking the sarcolemma, mitochondria, lysosomes, and nucleus. In CTR muscle, desmin was visible in the sarcolemma and cytoplasm with the expected banding pattern in longitudinal sections ( Figures 6A-C panels). In MUT muscle the presence of desmin appeared lower than in CTR muscle and its distribution pattern was different ( Figures 6D-F panels). The desmin banding pattern was absent in MUT muscle, and aggregates containing this protein were present at paracentral and peripheral locations within the muscle fibers ( Figure 6E).

CCT5, Alpha-B-Crystallin, Hsp90, and Desmin
The CS components CCT5, alpha-B-crystallin, and Hsp90, the latter two normally associated at the level of the Z-discs, and desmin were assessed to determine presence and distribution. We performed double immunofluorescence for: 1) desmin and CCT5 subunit ( Figure 7B); desmin and alpha-B-crystallin ( Figure 8C) desmin andHsp90 (Figure 9). Desmin was homogeneously distributed in CTR muscle along the Z-discs, seen as horizontal striations and with a striated pattern in cross-and longitudinal sections, respectively (green fluorescence in CTR panels of Figures 7-9). In MUT muscle, signal of desmin appeared slighter than in CTR muscle and appeared as irregular or trabecular patterns, and as dots (green fluorescence in MUT panels of Figures 7-9). In some fibers the desmin-labelled sarcolemma was not visible.
The cytoplasm of MUT muscle fibers showed little if any signal for CCT5 subunit, but this signal was present in the sarcolemma and the intercellular matrix, as also illustrated in Figure 3. Almost no colocalization of CCT5 with desmin was observed in the extracellular matrix of affected tissue (Figure 7, rightmost MUT panel).
Alpha-B-crystallin was uniformly distributed in CTR muscle showing a striated pattern in longitudinal and cross sections ( Figure 8). In contrast, in MUT muscle, alpha-B-crystallin signal was milder, appearing as a light band in longitudinal sections and as precipatetes with desmin in the sarcoplasm (white arrows in MUT merge panel of Figure 8). Likewise, colocalization of alpha-B-crystallin and desmin was reduced in half in MUT muscle as compared with CTR muscle.
Hsp90 was distributed throughout the cytosol of the CTR muscle fibers in a well-organized pattern in longitudinal and cross sections ( Figure 9, CTR panels). In the MUT muscle, Hsp90 was present but appeared in an irregular pattern somewhat like that of desmin ( Figure 9, MUT panels). Hsp90 and desmin colocalization was reduced in the MUT muscle as compared with the CTR muscle.

In Silico Analysis of the CCT5 Mutant Protein
The histopathological abnormalities found in the MUT muscle suggested poor protein homeostasis, which can, as a working hypothesis, be attributed to a defective CCT complex carrying the mutant CCT5 subunit. Therefore, we investigated the impact of the mutation on the CCT5 protein molecule. We used bioinformatics as FIGURE 3 | Localization of CCT5 on CTR and MUT muscles. Immunofluorescence was done with FITC conjugated anti-CCT5 antibody (green); nuclei were stained withe Hoechst (blue). Nuclear localization of CCT5 (white spots) in CTR muscle (18.09%) was more frequent than in MUT muscle (5.99%). Bar = 25 μm.
Frontiers in Molecular Biosciences | www.frontiersin.org June 2022 | Volume 9 | Article 887336 5 we did in a previous work with another mutant of CCT5 because, in that work, we detected molecular abnormalities caused by the mutation that accurately predicted the abnormalities of the CCT5 subunit molecular properties and functions that were also demonstrated by wet lab experiments (Min et al., 2014;Spigolon et al., 2017;Macario and Conway de Macario, 2020).

Comparison of Molecular Models
To learn about the impact of the mutation on the CCT5 apical domain configuration previously reported for the Leu224Val mutant (Antona et al., 2020), we superposed wild type and mutant molecular models under three conditions, nucleotide free, ATP bound, and ADP bound, Figure 10. It was confirmed that the mutant protein differs from the wild type, particularly at the level of the apical domain. FIGURE 4 | Double immunofluorescence for CCT5 and CCT1 subunits on CTR (top set of five panels) and MUT (bottom set of five panels) muscles. CTR and MUT muscles were stained with FITC conjugated anti-CCT1 antibody (green) and with TRITC conjugated anti-CCT5 antibody (red). Nuclei were stained with probe Hoechst (blue). Colocalization of CCT5 and CCT1 in CTR muscle (92.75%) and in MUT (89.07%) muscle appeared as white spots (see the two panels at the bottom left corner of the CTR and MUT sets of panels). Bar = 25 μm. Nuclear localization of CCT1 in CTR (5.63%) and in MUT (1.40%) muscles appeared as white spots (see the two middle panels at the bottom of the CTR and MUT series of panels with the legend "CCT1 nuclear colocalization"). Bar = 25 μm.

Radius of Gyration
We used the radius of gyration (RG) versus time method to determine if the distribution of atomic masses in the mutant was different from that of the wild-type protein under three conditions: nucleotide free, ATP bound, and ADP bound ( Figure 11). All trends of the mutant protein showed different RG values than the wild-type counterparts. For instance, the nucleotide-free mutant ( Figure 11, right panel, black) had a trend opposite to that of the wild-type nucleotide-free protein, with increasing values, 2.9-3.2 RG for the mutant and decreasing values, 2.9-2.7 RG, for the wild-type molecule ( Figure 11, left panel, black).

FIGURE 5 | Detection of myosin (MHC IIX) with FITC-conjugated antibody (green) and actin with TRICT-conjugated antibody (red) in CTR and MUT muscles.
Myosin positivity was higher in the MUT than in the CTR muscle whereas actin positivity showed the reverse pattern being higher in the CTR muscle. Also, the distribution patterns of the two proteins differed in the CTR and MUT muscles and colocalization was absent. Nuclei were stained with probe Hoechst 33342 (blue). Bar = 25 µm. The CCT5 molecule, like the other CCT subunits consists of three domains, apical, equatorial, and intermediate and while the latter two are composed of two separate segments of the protein's amino sequence, the apical domain consists of single segment (Supplementary Figure S2). Molecular dynamics simulations were carried out to determine if there were differences between the domain segments of the mutant and the wild type proteins nucleotide free, and ATP-or ADP bound. The RSMD analysis showed substantial differences between the two molecules, particularly at the level of the apical domain ( Figure 12).

Heat Maps
To identify the amino acids within each domain involved in the molecular conformational changes observed by molecular dynamics simulations, we resorted to heat maps. The heat maps show the RMSD value in Angstrӧm unit (Å, in y-axis) of each individual amino acid residue during simulations (frame in ns, in x-axis). Collectively, these analyses revealed instability of the apical domain of the mutant nucleotide-free protein ( Figure 13).

DISCUSSION
Many diseases of the central and peripheral nervous systems and of skeletal muscle are proteinopathies, namely, the central pathogenic-diagnostic molecule is a defective protein, which in most instances is the product of a mutant gene, thus the diseases are by definition genetic, and for the most part hereditary (Jameson et al., 2018). When the abnormal protein belongs to the chaperone system (CS), the disease can be classified as a chaperonopathy (Macario and Conway de Macario, 2005). In this report, we deal with a disease that according to clinical signs and symptoms is a genetic, early onset, distal motor neuropathy, associated to a mutation in a chaperone gene, the subunit CCT5 of the chaperone complex CCT (Antona et al., 2020). We report the histopathological characteristics of the skeletal muscle of the patient, focusing on details that in principle might be the result of deficient protein homeostasis caused by inadequate chaperoning. Information on the tissue abnormalities found in chaperonopathies and the molecular mechanisms underpinning these abnormalities, including the role of the defective chaperones, is scarce and badly needed (Macario and Conway de Macario, 2020). There is some information on the histopathological manifestations of myopathies, including one associated to a chaperonopathy, but little is available on specific aspects of protein homeostasis that might be affected by faulty chaperoning (Ruggieri et al., 2015;Cassandrini et al., 2017).
It is noteworthy that, in our patient, there is considerable disruption of the skeletal muscle architecture coinciding with abnormalities of desmin in the absence of cardiac manifestations of disease. Desmin is essential to maintain the structure of not only the skeletal but also the cardiac muscle. Diverse mutants of the desmin gene are associated with myopathy and cardiopathy (Brodehl et al., 2018). Alterations of desmin expression have been described in conditions of muscle atrophy induced by disuse and aging (Marzuca-Nassr et al., 2017). Also, the size of nuclei in muscle tissue may increase when desmin is functionally impaired (Heffler et al., 2020). If the desmin abnormalities in our patient are the result of deficient chaperoning by CCT, one may ask why the myocardium does not seem to be affected. An explanation could be derived from understanding the CS physiology. The CS components are distributed in all cells and tissues but assembled in groups of interacting molecules that are distinctive for each cell and tissue type, thus while the CCT chaperoning team may be present in the cytosol of all cell and tissue types its interacting partners (i.e., other teams to form networks, co-chaperones, chaperone co-factors, and chaperone interactors and receptors (Macario and Conway de Macario, 2020) are not the same in all locations. Therefore, a defective CCT team bearing a pathogenic mutant may show a range of capabilities depending on the cell type. This may explain the observation that desmin is considerably affected in the skeletal muscle while it is not in the cardiac muscle since the patient does not show clinical manifestations of heart disease. Another explanation could be that CCT does not participate directly in the folding of desmin, or in any of the steps leading to its maturation to become a functional protein in the appropriate cell locale but plays an indirect role. For example, CCT could be key for the folding and maturation of a protein, e.g., an enzyme, required by desmin production and migration to its physiological locations in skeletal but not in the cardiac muscle. If the CCT complex with the mutant CCT5 subunit fails in its chaperoning functions regarding that protein, desmin would be altered.
In the last few years, it has been found that, differently from other subunits, CCT5 (and in a less competent manner also the CCT4 subunit) has the ability to form homo-oligomeric complexes (micro-complex and single ring), supporting the hypothesis that single CCT5 homo-oligomeric rings may be the base assembly units for the formation of the functional hetero-oligomeric CCT hexadecamer (Sergeeva et al., 2019;Liu et al., 2021). A certain functional ability of the mutant CCT5 in our patient is suggested by its ability to form part of the CCT team, i.e., the CCT hexadecamer. This is suggested by the finding that CCT5 colocalized with CCT1, another component of the CCT team, in which these two subunits are not in direct contact (Leitner et al., 2012). Thus, one may conclude that if the mutant CCT5 colocalizes with CCT1 it must do so indirectly, through contact with the other subunits; and this would indicate that the mutant subunit must be competent to join the other subunits, FIGURE 8 | Double immunofluorescence with FITC-conjugated antibody for desmin (green) and with TRITC conjugated anti-alpha-B-crystallin antibody (red) in CTR and MUT muscles. Alpha-B-crystallin positivity in the sarcolemma and sarcoplasm of CTR muscle fibers appeared in the longitudinal (top row of CTR panels) and the transversal sections (middle row of CTR panels); this positivity was weaker in the MUT muscle fibers (bottom row, MUT panels). Fiber disorganization was evident (orange arrows on the MUT panel to the right, one before the last or "merge" panel). On this same "merge" panel, white arrows point to examples of alpha-Bcrystallin and desmin aggregates. Alpha-B-crystallin and desmin colocalization was higher in CTR muscle (longitudinal section, 83.57%; transversal section, 88.96%) than in MUT muscle (for example, only 46.89% in MUT muscle transversal section, as show in this figure). Nuclei were stained with probe Hoechst 33342 (blue). Bar = 25 µm.
Frontiers in Molecular Biosciences | www.frontiersin.org June 2022 | Volume 9 | Article 887336 9 form the octameric ring, and the full hexadecameric chaperoning machine. This is in line with the fact that the mutation affects primarily the apical domain of CCT5, namely, the domain involved in substrate recognition and chaperoning chamber closing and opening, but not in the formation of rings and hexadecamers (Lopez et al., 2015). Thus, one can expect a certain level of chaperoning ability from the CCT complex, only curtailed by insufficient capability for substrate FIGURE 9 | Double immunofluorescence with FITC-conjugated antibody for desmin (green) and with TRITC conjugated anti-Hsp90 antibody (red) in CTR and MUT muscles. Hsp90 positivity was well organized in longitudinal and transversal sections of the CTR muscle, but in contrast, it was disrupted and weaker in the MUT muscle (see for example the MUT muscle longitudinal fiber indicated by the orange arrow on the MUT "Merge" panel). Hsp90 and desmin colocalization was high in CTR muscle (longitudinal section, 84.51%; transversal section, 86.75%) by comparison with the MUT muscle (for example, it was 69.34% in MUT muscle transversal section, as shown in this figure). Nuclei were stained with probe Hoechst 33342 (blue). Bar = 25 µm. recognition and handling and depending on the cell type's proteome, which is different for each cell type. Consequently, one can expect diverse kinds and levels of failure of protein homeostasis in different cell and tissue types. To further complicate the landscape of deficient protein homeostasis in the whole organism stands the diversity of possible substrates for CCT, which include a variety of proteins: structural (Yam et al., 2008), regulatory of cell cycle and apoptosis (Willison, 2018a;2018b), and others involved in neural tissue development and degeneration (Chánez-Cárdenas and Vázquez-Contreras, 2012). Substrate selection and binding depend on specific amino acid sequences on the client protein (Willison, 2018a;2018b) and on the translational context (Yam et al., 2008). In different tissue types (with distinct proteomes), and under different conditions (e.g., normal vs stress condition), CCT can bind different substrates. This adds to the diversity of effects that a defective CCT complex may cause in various tissues, for example, striated skeletal and cardiac muscles, and nervous tissue and skeletal muscle.
Another finding that also indicates that the chaperonin complex with the mutant CCT5 subunit can function, at least to some extent and for a reduced range of substrates, is that the signal for actin, a canonical substrate for CCT (Berger et al., 2018;Willison, 2018aWillison, , 2018b, was only slightly affected in the patient's muscle. It can therefore be inferred that while actin, normally the substrate for which CCT has high avidity, is still recognized by the defective chaperone, whereas other less preferred substrates are not.  The RMSD values reveal differences between the two molecules and instability of the apical domain of the mutant. Similar trends were shown when ADP and ATP bound molecules were examined, although in the ATP-bound molecules the differences in instability of the mutant with regard to the wild-type molecule were less marked (data not shown).
Frontiers in Molecular Biosciences | www.frontiersin.org June 2022 | Volume 9 | Article 887336 The CCT complex, and the Hsp90 and alpha-B-crystallin chaperones are at the Z-disc of healthy skeletal muscle and are believed to play key roles in the development and maintenance of this structure (Roh et al., 2015;Unger et al., 2017;Berger et al., 2018). Our experiments demonstrate that all three components are affected in MUT muscle, which would impair the functionality of the Z-disc; Z-lines would not anchor the contractile filaments and, in turn, desmin would not transmit the force of contraction to sarcolemma, hampering the shortening of muscle fibers. Alpha-B-crystallin is a desmin chaperone but which of the two proteins is primarily damaged is to be investigated. Hsp90 and alpha-B-crystallin genetic variants are related to hereditary myopathies (Reilich et al., 2010;Unger et al., 2017). Many fibers of skeletal muscle tissue from the patient were in apoptosis, but TUNEL assay does not allow us to differentiate the classical apoptosis, from denervation and necrosis, and further investigations are required. However, a skeletal muscle histomorphology due to inactivity and comparable to the one described here is not reported in literature, whereas protein aggregation and reduction of myofibrils in zebrafish cct5-mutant model was accompanied be severe decrease of skeletal muscle-dependent force generation (Berger et al., 2018). Therefore, it is highly likely that failure of CCT entails defects involving alpha-B crystallin and Hsp90, which is not surprising considering the physiological interactions known to occur between different members of the CS.
The in silico analyses of the mutant CCT5 in comparison with the wild type counterpart, demonstrated changes in the mutant, most evident in the apical domain, which suggest that the chaperonin is defective and could be the cause of some of the abnormalities observed in our patient.
The CCT complex fold diverse substrates, typically actin and tubulin, which differ in structure. This suggests that substrates for CCT do not have to have a specific amino acid signature for recognition and binding by the chaperonin but must show other features for the purpose. For instance, matching tridimensional conformations in the chaperone and the substrate could be involved with participation of multiple weak interactions. In this regard, the radius of gyration and RMSD analyses revealed in the Frontiers in Molecular Biosciences | www.frontiersin.org June 2022 | Volume 9 | Article 887336 mutant molecule an energy disturbance, suggesting altered folding with opposite masses distribution and a different molecular equilibrium as compared with the wild type CCT5 subunit. The apical domain of the mutant subunit showed a major deviation of atoms in comparison with the wild-type counterpart, most notably under nucleotide-free conditions but also appreciable under ATPbound conditions. Helix-10 of the apical domain (amino acids 309 EANHLLLQ 316 ) and the proximal loop region (amino acids 234 FSHPQMPK 241 ) are involved in substrate-binding (Pereira et al., 2017). Beyond the substrate-binding region, the extended α-helix (helix-9) of the apical domain functions as a built-in lid to close the CCT hexadecamer's cavity and is linked to an apical loop (amino acids 259 KPKTKHK 265 ), which creates a highly charged region probably involved in substrate interaction (Pereira et al., 2017). Heat maps of the mutant CCT5 subunit confirmed the instability of the apical domain affecting helix-10, proximal loop, and helix-9 position in absence of nucleotide. It is, therefore, possible that the chaperoning functions of the CCT hexadecamer with the mutant CCT subunit are deficient with compromise of substrate recognition, binding, and conveyance. Thus, it is likely that at least some of the abnormalities observed in the striated muscle of the patient are the consequence of deficient chaperoning.
Here we report for the first time the histomorphology of a genetic chaperonopathy in the skeletal muscle tissue. We believe that, taking into account the difficulty to reproduce the complexity of a tissue in vitro, our results provide the indispensable platform for designing and launching the experiments necessary for elucidating the molecular mechanisms underpinning the reported tissue abnormalities.

DATA AVAILABILITY STATEMENT
The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding author.


Transcription Factor NRF2 Participates in Cell Cycle Progression at the Level of G1/S and Mitotic Checkpoints

Transcription factor NRF2 is a master regulator of the multiple cytoprotective responses that confer growth advantages on a cell. However, its participation in the mechanisms that govern the cell division cycle has not been explored in detail. In this study, we used several standard methods of synchronization of proliferating cells together with flow cytometry and monitored the participation of NRF2 along the cell cycle by the knockdown of its gene expression. We found that the NRF2 levels were highest at S phase entry, and lowest at mitosis. NRF2 depletion promoted both G1 and M arrest. Targeted transcriptomics analysis of cell cycle regulators showed that NRF2 depletion leads to changes in key cell cycle regulators, such as CDK2, TFDP1, CDK6, CDKN1A (p21), CDKN1B (p27), CCNG1, and RAD51. This study gives a new dimension to NRF2 effects, showing their implication in cell cycle progression.

Introduction
NRF2 (Nuclear factor (erythroid-derived 2)-like 2), encoded by the gene NFE2L2, is a transcription factor that belongs to the basic region leucine zipper (bZIP) family. It regulates the expression of over 250 genes through its heterodimerization with MAF proteins and interaction with its target sequence, the Antioxidant Response Element (ARE) [1]. NRF2 activates the transcription of cytoprotective genes in response to oxidative damage, electrophiles, xenobiotics, or pro-inflammatory stimuli, and is considered a master regulator of homeostatic processes, including cell detoxification, antioxidant cell defense, metabolism regulation, control of proteostasis, inflammation resolution, DNA repair, cell survival, etc. [2,3].
However, the impact of NRF2 on cell cycle progression and cell redistribution in the cell cycle phases has not been explored [16]. Cell transit throughout the G1, S, G2, and M cycle phases is governed by a protein regulatory network of cyclin-dependent kinases (CDKs) [17], whose activity is controlled by association with various phase-specific cyclins, regulatory phosphorylation, and binding of CDK inhibitors [18]. Together, these molecular engines propel cell proliferation through several checkpoints that ensure proper progression along the cell cycle according to mass cell growth and DNA replication, integrity, and division. In this study, we analyzed the effect of NRF2 throughout the cell cycle phases under standard proliferating conditions in cell cultures and identified correlations with the changes in key cell cycle regulators, such as CDK2, TFDP1, CDK6, CDKN1A (p21), CDKN1B (p27), CCNG1, and RAD51. This study gives a new layer to the characterization of the role of NRF2 in cell cycle progression. . For synchronization at G1/G0, cells were FBS-deprived for 64 h and then replated in a standard medium (0 h point). For synchronization before S entry, a double thymidine pulse was performed. Cells were treated with 2.5 mM thymidine (Abcam, ID ab143719, Cambridge, UK), and then washed with PBS twice and maintained in thymidine-free medium for 8 h. Then, a second thymidine pulse was performed under the same conditions, and cells were finally chased to standard medium (0 h point). For synchronization in the M phase, cells were treated with 75 ng/mL nocodazole (Sigma Aldrich, ID M1404) for 16 h. Then, round and partially detached cells were harvested and plated in a nocodazole-free medium (0 h point).

Analysis of mRNA Levels
Total RNA extraction and qRT-PCR were completed, as detailed in [13]. Primer sequences are shown in Table 1. Data were analyzed using the ∆∆CT method. Raw data were normalized using the geometric mean of the housekeeping genes GAPDH and TBP. All PCRs were performed from quadruplicate samples.

MTT and Cell Growth Assays
Cells were treated with thiazolyl blue tetrazolium bromide (MTT) (Sigma Aldrich, ID M5655) for 1 h, then the medium was retrieved, and the cells were directly lysed in DMSO for 30 min. One hundred µL of the supernatants were analyzed in 96-well multiwell plates at 550 nm in a VERSAmax microplate reader (Molecular Devices, San Jose, CA, USA). For cell growth kinetics, cells were counted in a Neubauer improved Bright-Line counting chamber (Marienfeld) using Trypan Blue solution (Sigma Aldrich, ID T8154) to discriminate the dead cells.

Flow Cytometry
For analysis of DNA content, the cells were harvested, pelleted, and fixed in cold 70% ethanol diluted in PBS for at least 24 h at −20 • C. Ethanol was then removed by centrifugation (1100 rpm, 5 min) and cells were washed twice with PBS containing 0.03% Triton-X100 (PBST) (Sigma Aldrich, ID 9002-93-1). Finally, cells were resuspended in PBST with 1µg/mL Hoechst 33342 (Thermo Fisher Scientific, ID H3570) and incubated for 30 min in darkness. Analysis of the DNA content was performed with a FACSCanto II (BD Biosciences) flow cytometer in at least 10,000 events per condition. Calculation of the number of cells in G1, S, and G2 was completed using the Modfit (version 5.0) software. For EdU analysis, the Click-iT™ EdU Alexa Fluor™ 647 Flow Cytometry Assay Kit (Molecular Probe, ID C10424) was used, according to the manufacturer's instructions. Briefly, cells were treated with 10 µM EdU for 1 h, and then harvested, fixed, and permeabilized. For analysis of DNA content, cells were counterstained with 1µg/mL Hoechst 33342 (Thermo Fisher Scientific, ID H3570) for 30 min in darkness. Analysis of EdU incorporation and DNA content was performed with a FACSCanto II (BD Biosciences) flow cytometer, analyzing at least 10,000 events per condition. For analysis of intracellular reactive oxygen species (ROS), cells were incubated for 1 h at 37 • C with 2 µM hydroethidine (HE) (Thermo Fisher Scientific, ID D11347) in the dark, then detached from the plate, washed once with cold PBS, and analyzed immediately. Oxidized HE fluorescence (Excitation/Emission: 518/606 nm) was analyzed in a CytoFLEX S Flow Cytometer (Beckman Coulter, Brea, CA, USA).

NRF2 Alters Cell Cycle Progression and Proliferation
To determine the relevance of NRF2 in cell cycle progression, we knocked down NRF2 in U-373 MG glioblastoma-derived cells ( Figure 1) and in two unrelated cell lines, U-87 MG (Figure S1A-C) and MDA-MB-231 ( Figure S1D-F), that yielded similar results. As shown in Figure 1A, after 3 days of interference, we confirmed a drastic reduction of NRF2 protein levels, and of its target HO-1, used as a control of NRF2 transcriptional activity. There was also a reduction in the G2/M phase marker Cyclin B. Flow cytometry analysis of cell cycle phase distribution was performed in the cells labeled with EdU, which identifies S-phase cells, together with Hoechst 33342, which gives a quantitative measurement of the DNA content ( Figure 1B,C). The quantification of EdU-negative cells indicated that the NRF2 knockdown increased G1/G0 and decreased the S and G2/M cell numbers. The analysis of the Hoechst 33342-stained cells without EdU staining, with the help of the Modfit software, yielded similar results ( Figure S2A,B) and was used in subsequent experiments to analyze the cell cycle phase distribution. Similar results were obtained with another shRNA for NRF2 knock-down ( Figure S2A-D), and with a dominant-negative NRF2 mutant lacking the transactivation domain ( Figure S2E,F) [21]. Together, these results indicate that the redistribution of the cells towards G1/G0 is a consequence of the NRF2 depletion. The cell redistribution in the G0/G1 and G2/M phases correlated with a change in the proliferation rate, as determined by cell counts ( Figure 1D) and metabolic activity ( Figure 1E) along 4 days of growth. This effect was also observed in the U-87 MG and MDA-MB-231 cells ( Figure S1C,F). In conclusion, the absence of NRF2 impairs cell cycle phase distribution and proliferation rate.
We next tested if chemical or genetic NRF2 activation might exert the opposite effect. We treated U-373 MG cells with the NRF2 activator sulforaphane (SFN) and, considering that NRF2 activators might have off-target effects, we also used a lentiviral vector encoding the highly active NRF2 ∆ETGE mutant, which is partially resistant to proteasomal degradation due to loss of interaction with its main repressor KEAP1 [1]. In both cases, we observed the expected increase in both the NRF2 and HO-1 protein levels concomitant with an increase in Cyclin B (Figure 2A), a reduction of G1/G0 cells, and an increase in G2/M cells ( Figure 2B,C). Together these results suggest that NRF2 propels cell cycle progression.

NRF2 Absence Delays G1 Exit
To determine if differences in the proliferation between shco and shNRF2 cells are due to a restriction in transit towards the S phase, we synchronized them in G1/G0 by 64 h of serum deprivation, followed by a change to serum-containing medium for cell cycle reentry. The NRF2 knock-down was confirmed by immunoblot analysis, and Cyclin B levels were used as a control of cell cycle progression ( Figure 3A). There was a slight oscillation in the NRF2 levels, with a modest increase along G1 that was maximal at the S phase, and a slight decrease at G2/M. According to the flow cytometry profiles ( Figure 3B,C), a notorious difference in cell cycle progression was observed after 24 h of serum stimulation.    In additional experiments, cells were synchronized with a double thymidine pulse and monitored for 9 h upon thymidine withdrawal ( Figure 4A). Although most of the cells stayed in G0/G1 after synchronization, the shNRF2 cells again showed a modest accumulation in the G2/M phase compared to the control shco cells, which was also observed in ( Figure 3B,C). Besides, there was not a clear difference in S phase entry and progression between the shco and shNRF2 cells. As shown in Figure 4B, the NRF2 protein levels increased after 2-4 h, as cells approached the G1/S transition and then slowly decreased as cells approached G2/M. The pentose phosphate pathway (PPP) provides NADPH and nucleotide precursors, both of which are required for anabolic pathways and DNA replication of proliferating cells [2]. Considering that NRF2 is a well-known activator of genes related to the oxidative and sugar interconverting phases of the PPP, we analyzed if NRF2-depletion could affect cell cycle progression. As shown in Figure 4C, NRF2 depletion led to a significant reduction of its canonical target HMOX1, encoding HO-1, and to a subtle decrease in the transcript levels of glucose 6-phosphate dehydrogenase (G6PD) and transketolase (TKT) that were at the limit of statistical significance. Therefore, NRF2-depletion had a modest impact on PPP downregulation.

The Absence of NRF2 Delays Mitosis Withdrawal
We analyzed if, as suspected from the results obtained with the double thymidine pulse, NRF2 depletion was affecting M exit. We synchronized the shco and shNRF2 cells in the M phase with nocodazole and then released them to analyze the M-to-G1 phase transit. The G2/M cell counts indicated that the shNRF2 cells exit mitosis more slowly than the shco cells ( Figure 5A,B). This observation was consistent with the reduced number of shNRF2 cells transiting throughout G1. Conversely, upon 2 h of nocodazole removal, 25.4% of shco vs. 20.1% in shNRF2 cells were in the G1 phase. This difference was maximal at 4 h (42.9% of shco cells vs. 33.7% of shNRF2 cells) and was still slightly observed by 10 h (51.6% of shco cells vs. 47.1% of shNRF2 cells). Consequently, NRF2 absence results in a delay in mitosis exit. In addition, we observed an increase in NRF2 protein levels after 2 h, concomitant with G1 entry, which reached plateau levels between 4 and 10 h ( Figure 5C). These results further support the increase in NRF2 protein levels as a requisite for G1/S transition.

NRF2 Absence Alters Regulatory Mechanisms of G1-S, Mitosis and DNA Damage Checkpoints
We analyzed the expression of a battery of genes involved in cell cycle regulation using a NRF2-knockdown approach and the RT 2 Profiler PCR Array Human Cell Cycle from Qiagen (see Materials and Methods). We grouped all genes with significant expression changes (p-value ≤ 0.05) by their biological function in a heat map ( Figure 6A). We detected a reduced expression of several G1/S progression genes (CDK2, TFDP1, CDK7, CKS1B, MNAT1, and SKP2), and upregulation of TFDP2 and CDK6. There was also an upregulation of CDKN3, CDKN1A, CDKN1B, and RB1, the negative regulators of G1 exit, and downregulation of CDC6, implied in S phase progression. In addition, there were also changes in the expression of mitosis checkpoint genes (CDC6, CDK7, CDKN3, CKS1B, CKS2, MAD2L1, MAD2L2, and MNAT1), and an upregulation of DNA damage checkpoint-related genes RAD1, RAD17, and CCNG1, and DNA repair RAD51. Then, we further analyzed the gene expression changes with a threshold p-value of 0.05 but with a restrictive fold of change of 1.5/0.66 ( Figure 6B). We still detected seven different genes with altered expression ( Figure 6B,C). This restrictive analysis suggested a need for NRF2 in the expression of positive regulators of G1-S transition (CDK2, TFDP1) and repression of negative regulators of G1-S transition (CDKN1A and CDKN1B). Importantly, NRF2 appears to be needed for optimal expression of DNA damage detection (CCNG1) and repair (RAD51). Some of these genes were further analyzed by immunoblot in U-373 MG, U-87 MG, and MDA-MB-231 cells ( Figure 6D), further confirming the upregulation of p27 (CDKN1B), p21 (CDKN1A), and the downregulation of CDK2 and RAD51 in the three cell lines. Altogether, these results provide additional information about the participation of NRF2 in controlling the regulation of G1/S and mitotic checkpoints. Considering that NRF2 is a master regulator of redox homeostasis, the changes in these cell cycle regulators might be redox-dependent. This hypothesis was tested in the presence of the pro-oxidant paraquat (PQ) (Figure 7). Both NRF2 absence and PQ treatment induced ROS accumulation ( Figure 7A,B). NRF2 depletion led to the expected increase in p21 or p27 levels, but the enhanced ROS accumulation induced by PQ did not further increase, or even decreased, p21 or p27 levels ( Figure 7C). By contrast, the RAD51 and CDK2 levels were reduced in shNRF2 cells and exhibited a further reduction in the presence of PQ. Consequently, changes in these cell cycle effectors in the absence of NRF2 seem to respond to different mechanisms, with RAD51 and CDK2 showing the best correlation with redox alterations.

Discussion
While most of the effects of NRF2 on proliferation have been analyzed in the presence of NRF2 activators and aimed at elucidating its participation in tumorigenesis, its participation in cell cycle progression was little explored. To address this issue, first, we analyzed the effect of NRF2 on cell cycle phase distribution and proliferation rate. Three different cell lines, U-373 MG, U-87 MG, and MDA-MB-231 yielded essentially similar responses to NRF2 depletion, increasing G1/G0 and decreasing G2/M cell counts, and slowing down proliferation. In other words, NRF2 depletion tends to stop cell cycle progression. These results are consistent with reports showing that the silencing of the NRF2 coding gene is associated with the induction of premature senescence [22].
NRF2 depletion led to an accumulation of G1 cells indicating that they cannot efficiently pass the G1/S restriction point. Besides, NRF2 levels seem to be maximal at S phase entry, suggesting that NRF2 deficiency prevents the proper preparation of cells to enter the S phase. This hypothesis is consistent with the observation that NRF2-deficiency led to a reduced expression of the positive cell cycle regulators CDK2 and TFDP1 and increased expression of the cell cycle inhibitors, CDKN1A and CDKN1B, encoding p21 waf and p27 kip1 , respectively.
The cell cycle kinase inhibitor p21 participates in cell cycle regulation, DNA repair, and oxidative stress response [23]. It has been reported that NRF2 transcriptionally activates p21 and promotes cell defenses against oxidative stress induced by hydrogen peroxide [24]. However, p21 also participates in the onset of cellular senescence, suggesting the need for a tight control regulation of its function [25]. p21 is an effector of the tumor suppressor protein p53, which is involved in the regulation of the cell cycle and DNA repair and is activated by oxidative stress [26,27]. P53 activation leads to transient expression of p21, triggering transient G1 cell cycle arrest [28]. Nevertheless, among the cell lines studied, U-373 MG cell line is known to have a mutated p53, suggesting that under our experimental conditions, p21 upregulation is p53 independent. Further work is required to determine the crosstalk among NRF2, p53, and p21 in the regulation of G1/S transition, but our results are consistent with a role of NRF2 in propelling cells through this checkpoint.
NRF2-knockdown also led to the upregulation of the CDK inhibitor p27 (CDKN1B). NRF2 deficiency has been reported to decrease the p27 levels in response to Angiotensin II in cardiomyocytes [29]. This effect was not due to the direct inhibition of CDKN1B and therefore it might represent a post-translational effect of Angiotensin II activity. In fact, our results are in line with [16], in which depletion of PERK1, a kinase that activates NRF2 under stress conditions, led to the downregulation of p27 and p21. It is most likely that the difference among these studies is related to the dual role of CDK inhibitors, stopping the cell cycle until cells are ready for S phase entry or leading to senescence if there is a persistent impediment [30,31].
NRF2-depletion also led to a reduced transit throughout G2/M. Our results are consistent with a previous study which reported that Nrf2-deficient primary epithelial cells exhibit redox alterations, and G2/M-phase arrest that could be partially reversed by treatment with N-acetylcysteine and glutathione [32]. We found a role of NRF2 in the control of DNA integrity. RAD51 participates in the homologous recombination and DNA repair, while CCNG1 is associated with G2/M phase arrest in response to DNA damage. Therefore, our findings are consistent with a deficiency in the surveillance of DNA integrity and timely repair, leading to the extension of G2/M duration. In agreement with our results, it has been previously reported that NRF2 facilitates the repair of radiation-induced DNA damage through the induction of RAD51 [8,33]. The authors identified putative NRF2-responsive enhancers in RAD51 and other repair genes that are involved in the homologous recombination repair pathway. However, our analysis of the Encyclopedia of DNA Elements (ENCODE) of the human genome failed to retrieve a significant site for NRF2 binding, or for its partner MAFK, to the promoter of RAD51 (data not shown).
Considering the need for NADPH in cycling cells for anabolic reactions and redox control, we explored the effect of NRF2-deficiency in the expression of two limiting genes of the oxidative and sugar converting phases of the pentose phosphate pathway (PPP), glucose 6 phosphate dehydrogenase (G6PD) [34], and transketolase (TKT) [35], which are regulated by NRF2 [2]. The transcript levels of G6PD and TKT were very little changed after NRF2 depletion, consistent with our previous results that the ROS levels of this cell line are low and insensitive to NRF2 modulation [13].
Nocodazole inhibits microtubule polymerization and therefore arrests cells at mitosis. Upon nocodazole withdrawal, the NRF2-depleted cells exhibited a delay in mitosis exit, suggesting that, even though NRF2 levels are low at mitosis, it exerts still unexplored effects on the transition through this cycle phase. However, we also detected reduced NRF2 levels in G2/M in cells synchronized with the double thymidine pulse. Then, considering the reduced NRF2 levels and chromatin condensation that would further limit its transcriptional activity, the role of NRF2 in this cell cycle phase may be transcriptionindependent. The transcription-independent effects of NRF2 have already been postulated in the widely used Nrf2-knockout mice, which expresses a fusion protein of N-terminal NRF2 bound to LacZ [36]. In these mice, NRF2 still serves as a protein tethering KEAP1, and, consequently, a reduction of NRF2 levels would lead to KEAP1 release for the degradation of other proteins [37]. The role of KEAP1 in the cell cycle has been reported for p21 in the G1/S transition [38,39] but its role in mitosis has not been reported yet. Considering that NRF2 depletion, and the consequent increased KEAP1 activity, leads to delayed mitosis, it is interesting to speculate that KEAP1 could somehow antagonize the proteolytic effect of Anaphase-Promoting Complex/Cyclosome (APC/C), which participates in the destruction of cycle regulators, including securing B type cyclins, to propel metaphase to anaphase transition [40].
NRF2 protein levels oscillate during cell cycle progression, being maximal at the G1/S, and minimal at the G2/M restriction points. NRF2 levels are regulated through a combination of modifications in its gene expression by specific transcription factors and epigenetics, as well as by modification of protein stability by the ubiquitin-proteasome system. Further work is required to identify the mechanisms that govern fluctuations in NRF2 levels and activity along the cell cycle.